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a Creative Research Initiative ‘Sousei’ (CRIS), Hokkaido University, N21W10, Kita-ku, Sapporo 001-0021, Japan
b Institute of Soil Science, University of Hohenheim, Emil-Wolff-Strasse 27, 70599 Stuttgart, Germany
c Institute of Plant Nutrition, University of Hohenheim, Fruwirthstrasse 20, 70599 Stuttgart, Germany
d Graduate School of Agriculture, Hokkaido University, N9W9, Kita-ku, Sapporo 060-8589, Japan
* Corresponding author (junw{at}chem.agr.hokudai.ac.jp)
Received for publication November 9, 2004.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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Abbreviations: DAT, days after transplanting • DGGE, denaturing gradient gel electrophoresis • PCR, polymerase chain reaction
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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The increase in CO2 concentration may influence plant growth and thereby plant–microbial interactions and nutrient cycling in ecosystems. Most experiments conducted with elevated CO2 concentrations showed a higher biomass gain of plants, grown at sufficient nutrient supply (Hodge and Millard, 1998), which can be attributed to a higher photosynthetic CO2 assimilation rate (Eamus, 2000). An increased allocation of 14C-labeled photosynthate (Rogers et al., 1994; Hodge and Millard, 1998) and increased carbohydrate concentrations (Norby, 1994) have been reported in roots at elevated CO2. This may be particularly important under conditions of nutrient deficiency, when a higher belowground allocation of carbohydrates promotes root growth for acquisition of nutrients. The increased input of C to roots could also stimulate root exudation of soluble organic compounds (Norby, 1994; Paterson et al., 1997; Cheng and Johnson, 1998), with putative effects on rhizosphere microbial populations, solubility of nutrients, and toxic elements. In this context, the nature of the exudates may be more important than their overall quantity (Cardon, 1996). Elevated CO2 concentrations may also affect the concentrations as well as the nature of some specific root exudates in the rhizosphere, which could have direct impact on nutrient acquisition and rhizosphere microbial activities. Alternatively, higher overall root exudation at elevated CO2 may be simply the consequence of a larger root system with unchanged exudation rates per unit root biomass or root length, and therefore unchanged rhizosphere concentrations.
In this study, white lupine was used as a model plant with extraordinary high expression of root-induced chemical changes, particularly under conditions of P limitation, and mainly confined to special bottle brush-like root structures, the so-called "cluster roots" (Neumann and Martinoia, 2002). Cluster roots are formed by a relatively small group of plant species such as white lupine and Proteaceae, which are adapted to habitats of extremely low soil fertility, usually without formation of mycorrhizal associations (Neumann and Martinoia, 2002). Cluster roots are the sites of intense exudation of organic metal chelators (organic acids, phenolics), protons and phosphatases, involved in mobilization of sparingly soluble inorganic and organic soil P sources (Gardner et al., 1983; Dinkelaker et al., 1989; Neumann et al., 2000; Wasaki et al., 2003). Therefore, white lupine provides a well-characterized model system to study the effects of elevated atmospheric CO2 concentrations on root exudation, the related changes in rhizosphere microbial communities, and nutrient availability in the rhizosphere.
In contrast to earlier studies dealing with CO2 effects on root exudation of white lupine in hydroponic culture (Watt and Evans, 1999; Campbell and Sage, 2002), we used a rhizobox soil-culture system with a P-deficient calcareous subsoil, with and without soluble P fertilization to characterize (i) CO2–induced alterations of root exudation and rhizosphere chemistry in different root zones, (ii) the potential impact on structural and functional diversity of rhizosphere microbial communities, and (iii) the potential impact on P availability and P acquisition in the rhizosphere.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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The rhizoboxes were fixed with a horizontal angle of 50° to stimulate root growth along the transparent root observation window of the boxes. Plant cultivation was performed under controlled environmental conditions in closed growth chambers with a 16 h day–8 h night cycle, light intensity of 300 µmol m–2 s–1, a 25°C day–20°C night temperature regime, a relative humidity of 60%, and either 400 µmol mol–1 (ambient) or 800 µmol mol–1 (elevated) atmospheric CO2 concentrations. Elevated CO2 concentrations were adjusted to 800 µmol mol–1 ± 5% by automatic injection of pure CO2 using an automatic CO2 controller (Siemens, Ditzingen, Germany) in one growth chamber. The ambient CO2 concentration of approximately 400 µmol mol–1, characteristic for the region of Stuttgart, was applied in the other growth chamber.
Plant Harvest, Collection of Root Exudates, and Soil Sampling
At 27 and 35 days after transplanting (DAT), filter paper discs (Model 2992, previously washed with methanol and deionized water; Schleicher & Schuell, Dassel, Germany), containing 60 µL cm–2 of deionized water, were placed onto the surface of lateral roots and cluster roots in different developmental stages (young, mature, and senescent clusters), appearing at root observation window of the rhizoboxes, for trapping organic acids released from the roots. The developmental stage of cluster roots was defined by color, length of the lateral rootlets, and distance from the root apex of the parent root, as described by Marschner et al. (2002). Filter discs with a size of 2.0 x 1.5 cm were used for exudate collection from mature and senescent root clusters and filter circles (5-mm diameter) for lateral roots and young cluster roots. Short-term collection (3 h) was performed to minimize microbial degradation of carboxylates and to recover a high proportion of root exudates (Neumann and Römheld, 2000). After 3 h of incubation, the filter papers and the corresponding roots with adhering soil were harvested separately into 1.5-mL microcentrifuge tubes. The surrounding rhizosphere soil (up to 3-mm distance from the root surface) was collected with a spatula and also transferred into microcentrifuge tubes. At the end of cultivation period, the root systems were washed out of the soil and the plants were separated into shoots and roots. All collected samples were frozen immediately with liquid nitrogen and stored at –20°C for further analysis.
Determination of Plant Dry Matter and Phosphorus Analysis
Dry matter production of shoot and roots was determined after oven-drying at 80°C for 2 d. Dried tissues were homogenized in a mill and each 0.25 g of the homogenized tissues were digested by dry-ashing at 500°C for 4 h. After cooling, the samples were extracted twice with 2 mL of 3.4 M HNO3 and evaporated to dryness for precipitation of SiO2. The ash was dissolved in 2 mL 4 M HCl, 10-fold diluted with hot water, and boiled for 2 min to convert meta- and pyro-phosphates to orthophosphate. Spectrophotometrical determination of orthophosphate was conducted according to the method of Gericke and Kurmis (1952).
Determination of Citrate in Root Exudates
Root exudation of citrate as major P-mobilizing carboxylate in white lupine, predominantly released from cluster roots (Neumann and Martinoia, 2002), was assessed by localized collection techniques (Marschner et al., 2002; Dinkelaker et al., 1997). Rhizosphere-soil solution containing root exudates was collected from different root zones of white lupine grown in rhizoboxes by use of filter papers with a high soaking capacity. Exudates trapped into filter paper were extracted with 1.0 mL deionized water for filter discs (2.0 x 1.5 cm) and 0.5 mL for filter circles (5-mm diameter). The citrate concentration in the water extracts was determined by an enzymatic method using a diagnostic test kit (R-Biopharm, Darmstadt, Germany).
Soil Enzyme Activities
The functional diversity of rhizosphere microbial communities and root secretion of extracellular enzymes was characterized by activities of marker enzymes involved in P (acid and alkaline phosphatases), N (chitinase, Leu- and Tyr-peptidases), and C (glucosidase, xylosidase, cellobiosidase) cycling in the rhizosphere (Marx et al., 2001; Kandeler et al., 2002). Acid and alkaline phosphatases, Leu- and Tyr-peptidases, chitinase, xylosidase, cellobiosidase, and glucosidase activities were measured with a fluorescence microplate reader (Flx800; Bio-Tec Instruments, Winooski, VT) using 4-methylumbelliferyl (MUF), phosphate, N-acetylglucosamide, xyloside, cellobioside, and glucoside as substrates for phosphatases, chitinase, xylosidase, cellobiosidase, and glucosidase, respectively. 7-Amino-4-methylcoumarin (AMC)-Leu and -Tyr were used as substrates for peptidase analysis. All substrates were dissolved with dimethylsulfoxide as a solvent and diluted to 1 mM with the appropriate buffer. MES buffer (0.1 M, pH 6.1) was used for the analysis acid phosphatase and enzymes involved in C cycling. To analyze the activities of alkaline phosphatase and peptidases, Trizma buffer (0.05 M, pH 7.8) was employed.
Rhizosphere soil was precisely weighed and suspended to 20 mg dry wt. mL–1 with sterile water. Twenty microliters of the soil suspension was mixed with 80 µL of the appropriate buffer and 100 µL of substrate solution in 96-well microplates (OptiPlate 96F; Greiner Bio-one, Frickenhausen, Germany). Fluorescence measurements were conducted for 3 h at 30°C with an excitation wavelength of 360 and 460 nm for fluorescence emission. Fluorescence readings were performed every 15 min. Results were expressed by the increasing rate of MUF and AMC liberation (nmol g–1 h–1).
Isoelectric Focusing and Activity Staining of Phosphatases
Rhizosphere soil around cluster roots (0.1 g fresh wt.) was transferred into a microcentrifuge tube. The soil was suspended by tapping with same weight of 0.1 M sodium phosphate buffer (pH 5.5). The soil suspension was incubated on ice for 5 min and occasionally mixed by tapping. After the incubation, the suspension was centrifuged at 15000 x g for 2 min at 4°C. Forty-seven microliters of supernatant were transferred into a new tube and mixed with 9 µL of 80% glycerol, 3 µL of 40% Bio-Lyte (pH 3–10; Bio-Rad Laboratories, Hercules, CA), and 1 µL of dye solution (56% glycerol, 0.5% bromophenol blue, 0.5% xylene cyanol). The mixed solution was used directly for electrophoretic separation, performed with a mini gel system (Mini-Protean III; Bio-Rad Laboratories) according to Ozawa et al. (1995).
After electrophoresis, the gel was transferred into 50 mL 0.1 M MES buffer (pH 6.1, for acid phosphatase) or 0.05 M Trizma buffer (pH 7.8, for alkaline phosphatase) and incubated 10 min with gently agitation. The equilibrated gel was soaked with substrate buffer (20 mM 4-methylumbelliferylphosphate in 0.1 M MES or 0.05 M Trizma buffer) for 5 min. The fluorescence of methylumbelliferone liberated by phosphatase activity was visualized under UV light (260 nm).
Polymerase Chain Reaction (PCR)–Denaturing Gradient Gel Electrophoresis (DGGE) Analysis for 16S rDNA
Changes in rhizosphere bacterial community structures were investigated by PCR–DGGE analysis for 16S rDNA (Marschner et al., 2002). DNA from bulk soil and rhizosphere soil samples was extracted using a Fast DNA Spin Kit for Soil (Qbiogene, Carsbad, CA) according to the manufacturer's instructions. After spectrophotometric determination of DNA concentrations at 260 nm, extracted DNA samples were stored at –20°C.
Bacterial 16S rDNA fragments were amplified using a primer set F984 and R1378 (Heuer et al., 1997). A gas chromatography (GC) clamp (Sheffield et al., 1989) was attached to F984 to improve the separation of the fragments. Twenty-five microliters of PCR reaction mixture contained 10 ng extracted DNA, 1 U Taq polymerase (Qbiogene), 1x PCR buffer with MgCl2 attached with the Taq polymerase, 0.16 mM each dNTP, and 2 pmol of each primer. A DNA thermalcycler (T3 Thermalcycler; Biometra, Göttingen, Germany) was employed to amplify the 16S rDNA specific fragments using the following program: 10 min at 94°C followed by 35 cycles of 1-min denaturation at 94°C, 1-min annealing at 55°C, and 2-min extension at 72°C followed by 10 min at 72°C and cooling at 4°C. Amplified fragments were confirmed by electrophoretic separation on a 2% agarose gel and purified using a QIAquick PCR purification kit (Qiagen, Hilden, Germany) according to the manufacturer's instructions.
Denaturing gradient gel electrophoresis was performed with 6% acrylamide gels and a linear gradient ranging from 35 to 55% (with 100% containing 7 M urea and 40% [v/v] formamide) using the Dcode system (Bio-Rad Laboratories). The acrylamide gel was polymerized overnight. Purified DNA samples were subjected to electrophoresis in 1x TAE buffer at 60°C at a constant voltage of 90 V for 16 h. Separated DNA fragments were detected with silver staining. The fragment patterns were analyzed using the Quantity One software (Bio-Rad Laboratories).
Statistical Analyses
Determinations of shoot and root biomass were performed with five replicates, whereas determination of enzyme activities, DGGE analysis, and analysis of root exudates were conducted with three replicates. Tables and figures show mean values and standard errors of the mean. Analysis of variance (p < 0.05) was performed using Microsoft Excel (Microsoft, 2001). Discriminant analysis based on enzyme activities was performed using SPSS Version 11 (SPSS, 2001).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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Citrate Exudation
Citrate was detected in all samples, but particularly high amounts were found in samples obtained from mature and even senescent root clusters (Fig. 1)
. Plants without P supply showed significantly increased citrate exudation in mature and senescent clusters. Elevation of CO2 did not significantly change citrate exudation independent of the P treatments.
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Activity distribution patterns of acid and alkaline phosphatases in the rhizosphere soil obtained from different root zones and cluster root developmental stages of white lupine were very similar (Fig. 2a–2d) . However, the absolute values of alkaline phosphatase activity in the different root zones were lower than those of acid phosphatase activity (Fig. 2a–2d). In the treatments without P supply, phosphatase activities were increased in all root segments, but particularly in senescent cluster roots. This effect was more expressed with increased duration of the culture period (Fig. 2b, 2d). There was a trend for increased phosphatase activities at elevated atmospheric CO2 concentrations (35 DAT), although differences were not significant in most cases (Fig. 2a–2d). Activity staining of acid phosphatase at pH 6.1 and of alkaline phosphatase at pH 7.1 after isoelectric focusing separation revealed one identical band with an isoelectric point of 4.7 for the different P and CO2 treatments (Fig. 3) .
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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Although elevated CO2 concentrations had no effects on total root biomass production, development of cluster roots was accelerated in both P treatments indicated by a higher proportion of older (mature and senescent) root clusters (Table 2). Accelerated development of cluster roots under elevated atmospheric CO2 concentrations has been similarly reported in hydroponic culture studies with white lupine (Watt and Evans, 1999; Campbell and Sage, 2002; Kania, 2004).
Cluster Root Activity
Despite cluster root formation under +P conditions (predominantly young clusters), citrate accumulation was preferentially confined to the rhizosphere of mature and even senescent root clusters of plants grown without P supply (Fig. 1). Increased citrate exudation from cluster roots is in line with the more severe expression of P limitation in the treatments without P application (Table 1), which has been similarly reported for white lupine in hydroponic studies (Neumann et al., 1999). However, the observation is somewhat contradictory to earlier findings obtained in hydroponic culture, where a pulse of citrate exudation was confined exclusively to mature root clusters and not to senescent cluster roots (Neumann et al., 1999; Watt and Evans, 1999). This may be attributed to citrate accumulation in the rhizosphere soil, which might be detectable even in later stages of cluster root development, such as senescent root clusters without secretory activity. On the other hand, prolonged longevity has been reported for cluster roots under soil conditions compared with plants grown in nutrient solution (Dinkelaker et al., 1995; Watt and Evans, 1999), which may increase also the period of secretory activity. Similar to earlier studies in hydroponic culture (Watt and Evans, 1999; Campbell and Sage, 2002; Kania, 2004), citrate exudation from individual root clusters in different stages of development was not significantly affected by elevated atmospheric CO2 concentrations (Fig. 1).
The activities of eight enzymes related to P, N, and C cycling in rhizosphere were measured in this study. Differential alterations of enzyme activities in the rhizosphere of different root zones were detectable only for chitinase and acid and alkaline phosphatases. Phosphatases are enzymes released from P-deficient plant roots (acid phosphatases) and microorganisms (acid and alkaline phosphatases) and are involved in mineralization of organic P forms in soils. Moreover, root secretory acid phosphatases can contribute to some extent to retrieval of organic P lost from roots into the rhizosphere (Neumann and Römheld, 2000).
The activities of both acid and alkaline phosphatases increased in the rhizosphere soil of white lupine, particularly in mature and senescent root clusters of the –P variants (Fig. 2), without significant effects of CO2 treatments. Accordingly, intense accumulation of acid phosphatase mRNA and protein and subsequent phosphatase secretion has been reported for roots of P-deficient white lupine in hydroponic culture, in response to P limitation (Ozawa et al., 1995; Wasaki et al., 1997, 2003), and particularly high phosphatase activities were released from mature and senescent root clusters with the lowest internal P status (Neumann et al., 1999, 2000). This is in line with the highest expression of phosphatase activity observed in the present study in the rhizosphere of plants without P supply, where shoot P concentrations indicate the most intense P limitation (Table 1). Therefore, acid phosphatase activity, predominantly detected in the rhizosphere soil of cluster roots in the –P variants, with increasing activity during the culture period (Fig. 2) may be mainly attributed to the root-secretory enzyme and not to phosphatases of microbial origin. Moreover, a very similar activity distribution pattern of alkaline and acid phosphatases in the rhizosphere soil of cluster roots (Fig. 2) and the appearance of only one identical band with an isoelectric point of 4.7 after isoelectric focusing separation with activity staining (Fig. 3), suggest that both acid and alkaline phosphatase activities are representing only one single enzyme. This may be explained by a very broad pH optimum reported for root-secretory acid phosphatase of white lupine (Tadano et al., 1993).
Similar to phosphatases, chitinase activity was particularly expressed in the rhizosphere of mature and senescent root clusters of P-deficient white lupine, although there was no clear relationship with the length of the culture period or CO2 treatments (Fig. 2). The preferential detection of chitinase in the rhizosphere of senescent root clusters is in accordance with differential expression of a chitinase gene in senescent cluster roots of P-deficient white lupine in hydroponic culture (Neumann et al., 2000), suggesting that similar to acid phosphatase, chitinase may be released from cluster roots into the rhizosphere. Chitinases produced by plant roots may play a role in antifungal plant–pathogen interactions but also in nondefensive functions, such as cell division, differentiation, and development (Collinge et al., 1993; Patil and Widholm, 1997). On the other hand, chitinase in soils can play a role in decay of organic N compounds and accordingly, depression of chitinase activity by readily available N fertilization has been reported (Olander and Vitousek, 2000).
Additional enzyme activities determined in the present study (Leu- and Tyr-peptidases, xylosidase, cellobiosidase, and glucosidase) did not show any clear relationship with different root zones, P fertilization, or atmospheric CO2 concentration (data not shown).
Soil Bacterial Community Structures
The results of PCR–DGGE analyses of 16S rDNA extracted from bulk soil and rhizosphere soil samples demonstrated that the developmental stage of individual root clusters had a larger influence on the bacterial community structures than the treatments of P and CO2 (Fig. 5 and 6). Similarly, Marschner et al. (2002) investigated the microbial community structure of cluster roots in white lupine grown in sand culture with soil microbial inoculation and concluded that the bacterial communities in the rhizosphere of cluster roots depended on plant age, the developmental stage of root clusters, and the related alterations in root exudation.
The relative abundance of DGGE bands was similar in bulk soil and in the rhizosphere of lateral roots but declined with increasing age of cluster roots, particularly expressed in mature and senescent root clusters (Fig. 6). This finding suggests that the diversity of bacterial populations declines probably as a result of specialization, related with the rapid root-induced changes in rhizosphere chemistry (pH, exudation of carboxylates and phenolics, redox potential, extracellular enzymes) during cluster root development (Neumann and Martinoia, 2002).
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CONCLUSIONS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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Contrasting reports on increased P deficiency–induced secretion of acid phosphatase (Moorhead and Linkins, 1997; Barrett et al., 1998) and citrate (Cardon, 1996) per unit root biomass in other plant species at elevated atmospheric CO2 concentrations may indicate a high variability of CO2 responses in root exudation for different plant species and/or culture conditions. Further experiments are also required to evaluate whether the response observed in white lupine is specific on cluster root plants.
Apart from short-term studies on CO2 effects at the level of individual plant species, also long-term cumulative effects of distinct plant C deposition via roots and litter fall and resulting effects on soil microbial communities and nutrient cycles need to be considered to evaluate effects of future elevated CO2 on a larger time scale.
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ACKNOWLEDGMENTS |
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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